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Abstract

Background— In diabetes or atherosclerosis, ectopic bone, fat, cartilage, and marrow often develop in arteries. However the mechanism is unknown. We have previously identified a subpopulation of vascular cells (calcifying vascular cells, CVC), derived by dilutional cloning of bovine aortic medial cells, and showed that they undergo osteoblastic differentiation and mineralization. We now show that CVC have the potential to differentiate along other mesenchymal lineages.

Methods and Results— To determine the multilineage potential of CVC, molecular and functional markers of multiple mesenchymal lineages were assessed. Chondrogenic potential of CVC was evidenced by expression of types II and IX collagen and cytochemical staining for Alcian blue. Leiomyogenic potential of CVC was evidenced by the expression of smooth muscle-α actin, calponin, caldesmon, and myosin heavy chain. Stromogenic potential of CVC was evidenced by the ability to support growth of colony-forming units of hematopoietic progenitor cells from human CD34+ umbilical cord blood cells for a period of 5 weeks. Adipogenic potential was not observed. CVC were immunopositive to antigens to CD29 and CD44 but not to CD14 or CD45, consistent with other mesenchymal stem cells. CVC retained multipotentiality despite passaging and expansion through more than 20 to 25 population triplings, indicating a capacity for self-renewal.

Conclusions— These results suggest that the artery wall contains cells that have the potential for multiple lineages similar to mesenchymal stem cells but with a unique differentiation repertoire.

Received March 4, 2003; de novo received June 11, 2003; accepted July 18, 2003.

In vascular disease, metaplastic or ectopic tissue elements often develop in the artery wall. Ectopic bone tissue in the artery wall includes complete trabecular architecture, Haversian canals, and osteoclast-like cells, as described for centuries.1,2 Ectopic cartilage, marrow, and fat tissue are also found in animal and human diseased vessels.1,3–5 The mechanism underlying this abnormal differentiation is not known. One possibility is that the ectopic tissue derives from pluripotent cells in the artery wall.

Vascular smooth muscle cells, well known to “de-differentiate” in vitro, also have phenotypic heterogeneity in vivo.6,7 Campbell and colleagues discovered different subpopulation phenotypes of smooth muscle cells that may correspond to in vivo phenotypic changes in wound healing.8,9 Vascular pericytes, counterparts to smooth muscle cells in the microvessels, have the capacity for osteoblastic differentiation in vitro.10,11 The lineages represented in ectopic arterial tissue resemble those produced by the embryonic neural crest, which is the origin of thoracic medial smooth muscle cells in the adult aorta.8,12 Bone morphogenetic proteins (BMPs), which regulate neural crest and mesenchymal lineage allocation,13 have been demonstrated in ectopic tissues of the artery wall.2,14

The existence of a mesenchymal stem cell in adult tissues was proposed by Caplan and Bruder.15 Mesenchymal cells are defined by their capacities for self-renewal and differentiation along various lineages. Normal and injured connective tissues appear to be constantly repopulated by immature mesenchymal cells derived from the marrow.16,17 Consistent with this, the neointimal cells of transplanted aortas were shown to derive from the recipient’s circulation.18,19 In vitro, marrow stromal cells can differentiate into bone, fat, cartilage, or muscle when treated with specialized induction and growth media.20 Cells with similar potential have been isolated from fat tissue21 and fetal calvaria22; these also require specialized induction media in order to differentiate.

We14,23,24 and others25,26 have shown that calcifying vascular cells (CVC), a subpopulation of cells from the artery wall and cardiac valves, have the ability to undergo osteoblastic differentiation and mineralization. It is not known whether such cells can differentiate along other mesenchymal lineages. To determine whether the artery wall contains multipotential mesenchymal stem cells, we tested CVC for lineage plasticity. Results showed that these cells have the capacity for chondrogenic, leiomyogenic (smooth muscle), and stromogenic (marrow stromal) lineages in addition to the osteogenic potential shown previously. Adipogenic potential was limited even with use of specialized induction media. These cells expressed the same surface CD antigens shown on marrow-derived mesenchymal stem cells,20,27 and they have substantial self-renewal capacity. These findings suggest that the artery wall contains mesenchymal stem cells with lineage plasticity and a unique differentiation repertoire.

Cell Culture

Bovine aortic smooth muscle cells (BASMC) were harvested, cultured, and passaged from explants, and CVC (used at passages 11 to 18) were subcultured from these cells by dilutional cloning, as described previously.14,23,24 This range of passages ensures that the features identified are retained despite passaging, avoids artifact from senescence, and confirms the cells’ capacity for self-renewal. CVC were grown in DMEM (Irvine Scientific) containing 15% heat-inactivated FBS (Hyclone Labs) and supplemented with sodium pyruvate (1 mmol/L), penicillin (100 U/mL), and streptomycin (100 U/mL), all from Irvine Scientific. Culture media were changed every 3 to 4 days until testing. Timing for each assay result is indicated in the corresponding figure.

For osteoblastic differentiation, medium was supplemented with 5 mmol/L β-glycerophosphate. This supplement is not required for osteoblastic differentiation in these cells, but it accelerates mineralization. No additional supplements were added for chondrogenic and leiomyogenic differentiation.

Alcian Blue and von Kossa Cytochemical Stainings

Cells were cultured at 40 000 cells per well in 24-well dishes. After 3 days or 11 days of culture, cells were washed with PBS, fixed, and stained with Alcian blue or von Kossa, using standard methods.

Alkaline Phosphatase Activity

Alkaline phosphatase activity was assayed as described previously.23 Briefly, we measured the activity of alkaline phosphatase in whole-cell extracts and normalized it for total protein content as determined by the Bradford method (Bio-Rad).

Western Analysis

Cells were cultured as described above, whole-cell extracts were prepared at indicated times, and Western analyses were performed.

Immunocytochemical Staining

Colony-Forming Unit Assay

Short-Term Culture

A hematopoietic progenitor cell colony formation assay was performed as described previously.28 Briefly, CVC or a positive control mouse marrow stromal cell line, S17,28 or BASMC were cocultured as a feeder layer with human hematopoietic cells for 10 days, and the number of hematopoietic progenitor cell colonies was assessed.

Long-Term Culture

CVC and a positive control mouse bone marrow stromal cell line,29 M2–10B4 (ATCC), were cultured separately in 6-well plates. At confluence, 1×104 human CD34+ umbilical cord blood (UCB) cells were overlaid on the CVC or M2–10B4 cells and cultured in 5 mL Iscove’s Modified Dulbecco’s Medium (IMDM) containing 10% FBS, and 10−4 mol/L β-mercaptoethanol. The cocultures were maintained for a period of 5 weeks. Conditioned media containing nonadherent cells were removed weekly, and coculture was refed with 2.5 mL of cell-free supernatant (conditioned media) and 2.5 mL of fresh media. After 1, 3, and 5 weeks, nonadherent cells were subcultured in triplicate in 24-well dishes with the use of MethoCult, a semisolid culture medium (methylcellulose media with agar). Colony-forming units (CFU), including granulocyte-macrophage colony-forming units (CFU-GM) and erythroid burst-forming units (BFU-e), were counted at day 14 under a light microscope (n=2).

Flow Cytometric Analysis

CVC were grown to confluence in 100-mm dishes and were harvested with the use of trypsin/EDTA; 106 cells were washed with ice-cold PBS and resuspended in 100 μL of PBS and transferred to flow cytometry tubes. The cells were incubated with primary antibody (≈15 μg/mL) at 4°C for 15 minutes, washed, and incubated with phycoerythrin-conjugated sheep anti-mouse secondary IgG antibody at 4°C for 15 minutes. The immunolabeled cells were washed and fixed, and flow cytometry was performed on a FACSCalibor with the use of CellQuest Software (BD Biosciences).

Time-Lapse Digital Videomicroscopy

Individual wells of CVC in 24-well plates were illuminated with a blue light–emitting diode at wavelength 470 nm. The short-wavelength monochromatic light was selected to reduce chromatic aberrations. Images were obtained with a ×10 objective, corrected for use at 75 mm, and projected onto a 0.33-inch monochrome CCD videocamera. Video frames were captured by an ATI Radeon Graphics board at a rate of 0.1 Hz with 640×480 pixel resolution. The images were compressed with the use of Motion JPEG, heximated, and set to play at 30 frames/s. Images were displayed in 640×480 format, with the diagonal dimension corresponding to ≈1.7 mm.

Results

To determine the multilineage potential of CVC, molecular and functional markers of chondrogenic, leiomyogenic, stromogenic, osteoblastic, and adipogenic lineages were assessed, and surface markers were identified by flow cytometry.

Chondrogenic Potential

To assess chondrogenic potential, whole-cell extracts were prepared from CVC cultures at the indicated times. Western analyses were performed to assess expression of type II collagen, a specific marker for chondroblasts. Results showed that type II collagen expression increased after confluence (Figure 1A). Another cartilage marker, type IX collagen, was also expressed in CVC at postconfluence (Figure 1B). Production of cartilage matrix acid mucopolysaccharides was assessed by Alcian blue cytochemical staining of CVC and noncloned BASMC cultures. CVC cultures were negative for Alcian blue staining at day 3 (Figure 1C, panel A) and positive at day 11 (Figure 1C, panel B, arrow). BASMC cultures were negative for Alcian blue staining even at day 11 (Figure 1C, panel C).

Leiomyogenic Potential

To assess the leiomyogenic potential of CVC, Western analysis for smooth muscle–specific proteins was performed with the use of whole-cell extracts from CVC and BASMC cultures at the indicated times. Results showed that smooth muscle-α actin was expressed at day 1 in both CVC and BASMC. The expression declined at day 4 in both cultures; however, smooth muscle-α actin expression increased from days 4 to 18 in CVC, returning to the level of expression at day 1. In contrast, in BASMC, expression continuously declined over time (Figure 2A), as described previously.7,30 A similar trend was observed with calponin expression in CVC, whereas its expression was minimal in BASMC (Figure 2A). Expression of smooth muscle myosin heavy chain (SM MHC) was also increased in a time-dependent manner in CVC, whereas its expression declined gradually in BASMC cultures (Figure 2A).

Figure 2. Leiomyogenic potential of CVC. A, Western blot analysis of whole-cell extracts from CVC and BASMC cultures, at the indicated times, using antibodies to smooth muscle α-actin (SM alpha-actin), calponin, and smooth muscle myosin heavy chain (SM-MHC) (n=3). B, Immunocytochemical staining for caldesmon (brown; 3,3′-diaminobenzidine stain counterstained with hematoxylin) in CVC (panels a and b) and BASMC (panels c and d) after 3 days in culture (n=3; magnification ×400). Panels b and d are controls, with secondary antibody used alone. C, Time-lapse digital videomicrographic frames 1 minute apart, showing contraction of CVC in upper left edge of the aggregate. Vertical line and arrow indicate position of cell front in upper panel. Cell contraction is toward the right. Shorter dimension in each panel is ≈500 μm (n=3).

To distinguish smooth muscle versus myofibroblastic differentiation, CVC and BASMC cultures were immunocytochemically stained for smooth muscle caldesmon after 3 days of culture. Both cell types were positively immunoreactive (Figure 2B). As a functional assay, contractility of CVC was assessed by time-lapse digital videomicroscopy at day 3 of culture. Spontaneous, coordinated contraction of groups of cells occurred at the edges of aggregates (Figure 2C).

Stromogenic Potential

One specific function of marrow stromal cells is to provide the environment required for hematopoietic cell growth. To assess the ability of CVC to differentiate along the marrow stromal cell lineage, we assessed their ability to support short- and long-term growth of hematopoietic cells. CVC were cocultured as a feeder layer with human hematopoietic cells separated by a layer of agar, and hematopoietic progenitor cell colony formation was assessed as previously described.28 After 10 days, the number of colonies supported by CVC was 41±36. By comparison, the positive control, S-17, supported twice as many colonies, and BASMC supported none.

To test CVC for longer-term stromogenic potential, we used a more stringent assay, as described previously.29 CVC and mouse marrow stromal cells (M2–10B4), as positive control, were grown as feeder layers for CD34+ human umbilical cord blood cells for a period of 5 weeks. Results showed that at the end of 5 weeks, CVC continued to maintain CFU at the same level as bona fide marrow stromal cells (Table). The conditioned media from CVC or M2-10B4 monocultures, a negative control, did not produce any CFU when cultured in MethoCult media (data not shown).

Total Colony-Forming Units Produced by Coculture of CD34+ Human Umbilical Cord Blood Cells With Either CVC or Positive-Control Marrow Stromal Cells (M2–10B4)

Osteoblastic Potential

We and others previously showed that CVC express markers specific for osteoblastic differentiation, including Cbfa-1 and osteocalcin.14,23,24 For the present study, the time course of osteoblastic differentiation was characterized by type I collagen expression by Western analysis, the time course of alkaline phosphatase activity, and the time course of mineralization. Results showed that type I collagen increased over the 8-day period (Figure 3A). Alkaline phosphatase activity, a specific marker for osteoblasts, showed increasing activity over a 2-week period (Figure 3B). By von Kossa staining, CVC were negative for calcium mineral at 3 days (Figure 3C, panel a) and positive at 11 days of culture (Figure 3C, panel b). In contrast, BASMC were negative for mineralization by von Kossa staining even at day 11 (Figure 3C, panel c).

Figure 3. Osteogenic potential of CVC. A, Western blot analysis of whole-cell extracts from CVC cultures at the indicated days in culture, using anti–type I collagen antibody (n=4). B, Time course of alkaline phosphatase activity through day 14 (n=4). C, Von Kossa staining of CVC cultures at day 3 (panel a) and day 11 (panel b) and of BASMC culture at day 11 (panel c) (n=3; magnification ×40).

Adipogenic Potential

To assess the adipogenic potential of CVC, cells were cultured in adipogenic induction media and formation of adipocytes were assessed by oil red O staining. Despite use of a variety of different induction media,20,21 oil red O staining was negative for up to 2 weeks of culture, with the exception of 2 instances out of 10 tests, in which a few oil red O–positive cells were observed. Insignificant levels of PPAR-γ2 were seen on Western blot analysis (data not shown).

Self-Replication

CVC replicated as undifferentiated cells for ≈20 to 25 passages, with retention of multipotentiality. These cultures are derived from single-cell clones, obtained by dilutional cloning from explants of bovine aortic medial tissue as described previously.14,23 Cells of each lineage survived for at least 3 to 5 weeks.

Discussion

The present findings indicate that the adult artery wall contains cells with lineage plasticity and self-renewal capacity. These cells, previously termed calcifying vascular cells (CVC), are thus a type of mesenchymal progenitor cell of the artery wall, with one distinction from mesenchymal stem cells. The apparent lack of adipogenic lineage in their differentiation repertoire suggests that CVC represent a stage of commitment one generation below the mesenchymal stem cell in the mesengenic lineage hierarchy.

Marrow-derived mesenchymal stem cells have been characterized by flow cytometry as positive for CD29 and CD44 and negative for CD45 and CD14.20,27 Our results showed that CVC express the same set of surface CD antigens that have been shown to express in mesenchymal stem cells, suggesting that CVC share a surface marker profile with known mesenchymal stem cells.

Interestingly, the time course of leiomyogenic differentiation in CVC differs from that of BASMC and other vascular medial cells. In phenotypic modulation, vascular medial cells lose expression of contractile proteins continuously from day 1 as they dedifferentiate to a synthetic state.7,30 Steitz et al31 found that SMC lose their contractile markers simultaneously as they gain osteoblastic markers in a time-dependent manner. However, Proudfoot et al32 showed that the expression of these markers varies with location, being greater in nodule-forming cells than in the surrounding monolayer of SMC, raising the possibility of different time courses among different subpopulations. Our present results show that both contractile markers and osteoblastic markers increase simultaneously in CVC, the nodule-forming subpopulation of BASMC. This finding is consistent with those of previous reports,31,32 and it suggests that in conventional SMC cultures, the increase in contractile markers in the CVC subpopulation may be masked by the loss of these markers by the remaining cells, which make up the majority of the culture. Conversely, the expression of multiple lineage markers by BASMC cultures30–32 may be attributable in part to heterogeneity33 and the presence of the CVC subpopulation.

It is not clear whether simultaneous actin expression and osteoblastic differentiation in CVC serves a function. One possibility is that these cells may require a 3-dimensional matrix environment for differentiation, and actin may be necessary for contraction of cells into nodular structures that provide this microenvironment. Schor and colleagues10 previously showed that pericytes form nodules by coordinated contraction of the aggregating cells. Our time-lapse analysis shows the same process in CVC. Previous investigators have also found that single-cell–derived mesenchymal stem cell cultures can undergo differentiation along multiple lineages within a single culture.34

Although these CVC differentiate along four other lineages, adipogenic potential could not be induced, even with addition of various adipogenic induction media. It remains possible that these cells have adipogenic potential but that the appropriate induction media or culture conditions have not been identified. On the other hand, if the lack of adipogenesis reflects an intrinsic lineage limitation of these cells, it is difficult to explain the presence of ectopic fat tissue in atherosclerotic plaque, unless cells with adipogenic potential arrive from the circulation or migrate in as pericytes from angiogenic vessels. Caplice et al35 and Hirschi and Goodell36 have provided evidence that marrow stromal cells may be the origin of pluripotent progenitor cells in connective tissues, such as CVC.

CVC have intriguing relations to marrow stromal cells (MSC). To our knowledge, no adult mesenchymal cells other than CVC and marrow stromal cells support hematopoietic cell growth in long-term culture assays. Of particular interest is that the mesenchymal stem cells derived from MSC are located in the marrow vasculature.37

The lack of apparent adipogenic potential suggests that CVC may represent second-generation pluripotent cells that are intermediate between mesenchymal stem cells and terminally differentiated mesenchymal cells. Given the potential use of stem cells for cellular transplant therapy, these cells may serve as an alternate source in therapeutic circumstances in which adipogenesis would be undesirable.

Acknowledgments

This study was supported by National Institutes of Health grants HL-30568 and HL/RA 69261; the Laubisch Fund; the Stanley J. Sarnoff Endowment for Cardiovascular Science, Inc (to K. Radcliff); the Cohen Fund; and the Guthman Fund. We thank Dr J. Fraser (UCLA) for the human umbilical cord blood cells and the use of FACSCalibor and CellQuest Software; Drs Alan Garfinkel, Robb MacLellan, and John Parker (UCLA) for assistance with time-lapse digital videomicrography; and Dr H. Clarke Anderson, Department of Pathology and Laboratory Medicine, University of Kansas, for reviewing the manuscript.